Chapter 5 Multiple Approaches to the Study of Chemokine Receptor Homo‐ and Heterodimerization

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Multiple Approaches to the Study of Chemokine Receptor Homo- and Heterodimerization Jose´ Miguel Rodrı´guez-Frade, Laura Martinez Mun˜oz, and Mario Mellado Contents 1. Introduction 2. Biochemical Techniques to Measure Chemokine Receptor Oligomerization 2.1. Western blot and immunoprecipitation 2.2. Colocalization assays 2.3. Fluorescence labeling of antibodies 2.4. Construction of fluorescence-labeled receptors 3. Resonance Energy Transfer (RET) Techniques 3.1. Bioluminiscence resonance energy transfer (BRET) techniques 3.2. Fluorescent resonance energy transfer (FRET) techniques 4. Sequential BRET-FRET (SRET) Technology 5. Conclusion Acknowledgments References

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Abstract Chemokines belong to a family of structurally related chemoattractant proteins that bind to specific seven-transmembrane receptors linked to G proteins. They are implicated in a variety of biologic responses ranging from cell polarization, movement, immune and inflammatory responses, as well as prevention of HIV-1 infection and cancer metastasis. Recent evidence indicates that chemokine receptors can adopt several conformations at the cell membrane. Chemokine receptor homo- and heterodimers preexist on the cell surface, even in the absence of ligands. Chemokine binding stabilizes specific receptor conformations and activates distinct signaling cascades. Analysis of the conformations

Department of Immunology and Oncology, Centro Nacional de Biotecnologı´a/CSIC, Madrid, Spain Methods in Enzymology, Volume 461 ISSN 0076-6879, DOI: 10.1016/S0076-6879(09)05405-6

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2009 Elsevier Inc. All rights reserved.

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adopted by the receptors at the membrane and their dynamics is crucial for a complete understanding of the function of these inflammatory mediators. We focus here on conventional biochemical and genetic methods, as well as on new imaging techniques such as those based on resonance energy transfer, discussing their advantages, disadvantages, and possible complementarity in the analysis of chemokine receptor dimerization.

1. Introduction The family of low molecular weight proinflammatory cytokines termed chemokines were originally described as specific mediators of leukocyte directional movement (Mackay, 2001). Current views nonetheless implicate these molecules in the movement of several cell types, because they participate in functions such as lymphocyte trafficking (Baggiolini, 1998), regulation of T cell differentiation (Sallusto et al., 1998), HIV-1 infection (Berger et al., 1999), angiogenesis (Belpario et al., 2000), development (Raz, 2003), and tumor metastasis (Mu¨ller et al., 2001). Today, scientists refer to the nearly 50 known chemokines as either constitutive chemokines, which are usually regulated during development, or as inducible chemokines, whose expression is regulated mainly by inflammatory mediators (Proudfoot, 2002). In addition, certain viruses encode highly selective chemokine receptor ligands that can serve as agonists or antagonists and may, thus, have a role in viral dissemination or evasion of host immune response (Alcami, 2003). On the basis of their broad range of functions, it is easy to deduce that chemokines must be central to a variety of diseases that are characterized by inflammation and cell infiltration. They have become a major focus of interest as therapeutic targets, because there is a clear correlation between the expression of specific chemokines and the orchestrated recruitment of cell populations during the course of some disease processes (Proudfoot, 2002). The chemokines act by binding to class A rhodopsin–like, seventransmembrane G-protein–coupled receptors (GPCR) (Horuk, 2001). The 20 receptors characterized to date are classified as CCR, CXCR, CX3CR, and XCR on the basis of their ligand specificity (Rossi and Zlotnik, 2000). Another group of receptors (D6, DARC, and CCXCKR) that can interact with several chemokines were recently denominated ‘‘silent’’ receptors, because they are unable to activate signal transduction events that lead to cell chemoattraction (Borroni et al., 2008). Most chemokine receptors are able to interact with more than one chemokine (shared receptors), although there are some examples of specific chemokine-receptor pairs (specific receptors) (Horuk, 2001). Expression of these receptors is finely regulated by factors that include cytokines,

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growth factors, and cell cycle status (Loetscher et al., 1996; Papadopoulus et al., 1999; Parks et al., 1998). It is, therefore, not surprising that cells respond differently to a chemokine, depending on the microenvironment in which they are found. As the chemokine receptors integrate numerous signaling pathways (Soriano et al., 2003; Thelen and Stein, 2008), the chemokine-mediated signaling cascade is more complex than was originally thought. Although initially considered a cytokine habit (Thelen and Baggiolini, 2001), various studies have demonstrated the existence of chemokine receptor homo- and heterodimers and have speculated on the functional relevance of these conformations (Mellado et al., 2001a,b; Percherancier et al., 2005; Vila-Coro et al., 2000; Wang et al., 2006). The difficulties in detecting these complexes with immunoprecipitation methods suggest conformational instability in the absence of ligand, but resonance energy techniques clearly show that chemokine receptor dimers form spontaneously in the absence of ligand (Hernanz-Falco´n et al., 2004; Wilson et al., 2005). A number of questions remain to be answered, however, including the dynamic nature of these receptor complexes and the role of distinct ligands in promoting the conformational changes that trigger function. This is particularly important in the case of chemokines, because there is a relative lack of selectivity in ligand binding, with many receptors showing high affinity for more than one chemokine (Tian et al., 2004) and simultaneous expression of several receptors on the same cell. Although biochemical technologies were classically used to analyze protein-protein interactions, our current knowledge has been supplemented by new approaches on the basis of energy transfer between fluorochromes followed by confocal microscopy. These methods exploit important technologic advances such as laser light sources, fluorescent probes, and improvements in computer science that allow digital imaging and image analysis. Independently of the technique used, the cell system being used must be characterized in detail before attempting analysis of chemokine receptor dimerization. This includes routine testing such as analysis of cell cycle status, cell surface receptor expression, and determination of receptor number and affinity constants, especially when cells are transfected with mutant or fluorescently labeled receptors.

2. Biochemical Techniques to Measure Chemokine Receptor Oligomerization Until recently, most assays used to demonstrate receptor oligomerization were based on biochemical approaches such as immunoprecipitation or crosslinking. Alternately, dominant negative receptor mutants were used to

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abrogate wild-type receptor functions by forming nonfunctional complexes (Rodriguez-Frade et al., 1999). Because any single method alone has intrinsic limitations, most should be used in concert with others to obtain meaningful results.

2.1. Western blot and immunoprecipitation Although the general protocol for immunoprecipitation assays is very similar, lysis buffer compositions vary, and cell number should be adjusted for each case. Cells (1
107 cells/ml), unstimulated or stimulated with the appropriate chemokine, are diluted immediately with 1 ml cold PBS to terminate stimulation. Cells are centrifuged (800g, 5 min, 4  C), washed with cold PBS, resuspended in 200 ml lysis buffer (10 mM triethanolamine, pH 8, 150 mM NaCl, 1 nM EDTA, 10% glycerol, 2% digitonin), and incubated (30 min, 4  C, with continuous rocking). Other lysis buffers can be also used; however, detergents can alter receptor interactions and must be evaluated carefully. A preclearing step is essential to reduce nonspecific binding to the immunoprecipitating antibody (Ab). For this step, incubate the supernatant containing solubilized proteins with antiimmunoglobulin Ab (antibody against immunoglobulin of the animal species from which the immunoprecipitating Ab is derived) coupled to agarose and incubate (30 min, 4  C). Then add the immunoprecipitating Ab (90 min, 4  C), followed by anti-Ig coupled to agarose (without washing; 60 min, 4  C). After extensive washing and centrifugation, resolve the pellets by SDS-PAGE. To adjust the percentage of the acrylamide solution, remember that the predicted molecular weight of chemokine receptors is in the 30- to 50-kDa range. Transfer the gel to nitrocellulose membranes and develop Western blot as described elsewhere (Rodriguez-Frade et al., 1999). The immunoprecipitation technique is based on the use of chemokine receptor-specific Ab and depends greatly on their characteristics (specificity, affinity). This method is useful for evaluation of receptor heterodimers, in which case one receptor should appear in the immunoprecipitates of the other receptor. Alternately, it can be applied to analyze homodimerization, although in this case the receptors must be tagged appropriately. Receptor labeling bypasses the need to raise antibodies specific for target receptors and has been used successfully to demonstrate homodimerization of CCR2 and in the case of other GPCR, such as b2Ars, GABAB, mGluR5, d-opioid, m3-muscarinic, and Ca2þ receptors (Angers et al., 2002; Rodriguez-Frade et al., 1999). Selection of an appropriate tag and its location in the receptor are both important factors. Amino acids added to the extracellular region might alter ligand binding, whereas modification of intracellular domains can disturb the coupling of signaling molecules. Both ligand affinity and receptor distribution should first be evaluated to ensure that the behavior of the tagged receptor is similar to that of the wild-type receptor. FLAG, myc,

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HA, GST, His, GFP, and GAL are among the tags for which there are commercially available antibodies that immunoprecipitate and recognize the tagged receptor in Western blot. Homemade epitopes can also be designed and included into the receptor cDNA sequence with standard PCR techniques (Rodriguez-Frade et al., 1999). Immunoprecipitation methods can be complemented with crosslinking assays with bifunctional reagents such as disuccinimidyl suberate (DSS). Before lysis, cells should be resuspended in 1 ml cold PBS with 10 ml 100 mM DSS and incubated (10 min, 4  C). Care must be taken in cell handling before the lysis step, because the presence of nonintact cells increases the background of nonspecific protein crosslinking. Prepare the DSS reagent just before use. Continue with lysis, immunoprecipitation, and Western blot analysis as described previously. In this case, Western blot analysis with specific Ab will develop the band corresponding to the monomeric receptor, as well as the high molecular weight dimeric, trimeric and oligomeric species.

2.2. Colocalization assays Modern optical microscopy allows us not only to visualize organelles and molecules but also to study their function. In living cells, we can analyze how a molecule moves, changes location, or associates with other molecules. Such phenomena were originally evaluated with colocalization assays, which detect light from two different fluorophores and evaluate a digital image for the presence of the same pixel in two distinct channels. Signal colocalization indicates adjacency of fluorophores, and thus of the molecules they label (Fig. 5.1). A high-numerical aperture microscope lens permits resolution near 300 nm, sufficient to locate molecules in different cell compartments but not to demonstrate molecular association. The technique requires fluorescence-labeled antibodies or receptors coupled to fluorescent proteins. To determine colocalization between chemokine receptors, plate cells on coverslips coated with poly-L-lysine (20 mg/ml, 1 h, 37  C) and culture them (24 h, 37  C, 5% CO2). After washing, fix the cells with 4% paraformaldehyde (10 min, room temperature [RT]). To avoid nonspecific binding, treat the cells with PBS supplemented with 1% BSA, 0.1% goat serum and 50 mM NaCl (1 h, 37  C). Add the receptor-specific Ab (30 min, RT). To facilitate the procedure, use antibodies of distinct species origin (i.e., mouse, rabbit, rat, hamster) to stain the two receptors. If prelabeled Abs are available, their use precludes the need for secondary antibodies. Otherwise, add the mixture of prelabeled secondary Ab (20 min, RT). If both primary Abs are of the same origin and isotype, add one of these Abs, followed by its fluorochrome-labeled secondary antibody. Wash and incubate the cells with undiluted serum from the same species as the primary Ab (30 min, RT). Now stain the cells with the specific Ab for the second receptor as

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A Cy

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Figure 5.1 Chemokine receptors colocalize at the cell membrane.(A) Scheme of the colocalization experiment. Chemokine receptors can be fused to fluorescent proteins (left) or immunostained with fluorochrome-labeled specific antibodies (right). (B) In a representative experiment, CCR2/CCR5 stably transfected L1.2 cells were fixed and costained with anti-CCR2 (anti-CCR2-Cy3) and -CCR5 mAb (anti-CCR5-Cy2).The merged image is also shown; arrows indicate colocalization areas (yellow). All images are overlaid on the DIC (differential interference contrast) image.

previously, followed by its secondary antibody labeled with a different fluorochrome (20 min, RT). Evaluate fluorescence on a confocal microscope with filters appropriate for the fluorochromes used.

2.3. Fluorescence labeling of antibodies Although commercially available Ab can be used for these purposes, antibodies can also be labeled in the laboratory. Given their intense fluorescence and low hydrophobicity, the Cy dyes are efficient tags for fluorescence labeling. In the standard labeling procedure, the contents of a commercial vial (‘‘to label 1 mg of protein’’) of Cy2, Cy3, or Cy5 are dissolved in 50 ml dimethylsulfoxide (DMSO). The antibody is dissolved to 1 mg/ml in buffer (100 mM NaCl and 35 mM H3BO3, pH 8.3). Mix 10 ml of dye/DMSO mixture with 200 ml antibody solution and incubate (30 min, 25  C, in the dark). Separate unbound dye by adding 300 ml of 100 mM NaH2PO4, incubate (30 min, 25  C), and load the sample on a PD-10 column preequilibrated with 100 mM NaCl, 50 mM NaH2PO4, 1 mM EDTA

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(pH 7.5). Elute the labeled protein with 2 ml of distilled H2O. The labeled antibody should be titrated against the chemokine receptor before use.

2.4. Construction of fluorescence-labeled receptors For rapid, easy visualization of chemokine receptors, fluorescent proteinbased constructs and microscopy techniques are very helpful. The constructs are typically used in colocalization analysis and energy transfer techniques. Receptors are cloned with standard molecular biology methods into commercially available vectors bearing fluorescent proteins. Insertion of the fluorescent probe in the C-terminal region of the receptor involves elimination of the receptor stop codon, whereas insertion in the N-terminal region requires elimination of the fluorescent protein stop codon. Transfected cells should be analyzed for receptor expression and function.

3. Resonance Energy Transfer (RET) Techniques Newer methods to determine chemokine receptor oligomerization are based on resonance energy transfer (RET). These techniques are also useful for determining conformation dynamics, the role of ligand and receptor levels, and for defining the dimerization site within the cell (Harrison and van der Graaf, 2006). There are two main types of RET, bioluminescence resonance energy transfer (BRET) and fluorescence resonance energy transfer (FRET). In the former, the donor molecule is luminescent (Pfleger and Eidne, 2006); in the latter, the donor fluorochrome transfers energy to an acceptor fluorochromes (Cardullo, 2007). Both techniques require generation of fusion proteins between the receptor and the fluorescent/luminescent donor and acceptor proteins, as well as the use of transfected cells (Boute et al., 2002). Controls must, therefore, be included to rule out alterations in receptor distribution between the cells and to avoid differences in receptor-mediated cell function. Although BRET has been used at the single cell level (Coulon et al., 2008), it is, in fact, an approach for cell suspensions (Pfleger et al., 2006). It allows measurement of energy transfer between receptors independently of their expression pattern and permits quantitation. In contrast, FRET imaging techniques use confocal or wide-field microscopy, allowing measurements in single cells and identification of cell locations at which FRET is detected.

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3.1. Bioluminiscence resonance energy transfer (BRET) techniques BRET makes use of nonradiative energy transfer between a light donor and a fluorescent acceptor. The bioluminescent energy resulting from the catalytic degradation of a substrate by luciferase is transferred to an acceptor fluorophore, which in turn emits a fluorescent signal (McVey et al., 2001). This transfer takes place when there is an effective range of 10 nm between donor and acceptor, which allows at least 50% of the energy to excite the acceptor molecule. For maximum spectral overlap, the acceptor fluorophore (YFP or GFP2) varies depending on the substrate oxidized (coelenterazine or its derivative DeepBlueC, respectively). Because of their small size and their hydrophobicity, coelenterazines cross the cell membrane easily. For BRET measurements, at 48 h posttransfection, wash cells once with PBS. Add coelenterazine H (Nanolight Technology) to a final concentration of 5 mM in PBS, and take readings with a multidetector plate reader that allows the sequential integration of signals detected in the 480  20 nm and 530  20 nm windows for luciferase and YFP light emission, respectively. The BRET signal is determined by calculating the ratio of the light intensity emitted by receptor-YFP to the light intensity emitted by receptor-RLuc. The values are corrected by subtracting the background BRET signal as measured in the same cells expressing the receptor-RLuc or the receptorYFP construct alone. For acquisition of full BRET spectra, cells are transfected with different amounts of receptor-YFP for a given quantity of receptor-RLuc. Cells are detached and resuspended in HBSS containing 0.1% (w/v) glucose. Cells (2
105) expressing different acceptor/donor (YFP/RLuc) ratios are seeded in 100 ml HBSS in a clear-bottom 96-well plate, and a BRET scan is performed by reading luminescence between 400 and 600 nm, immediately after coelenterazine addition. YFP fluorescence is determined in the same cells with a black 96-well plate. For BRET titration experiments, net BRET ratios are expressed as a function of the acceptor/donor ratio. These BRET saturation curves provide an idea of the maximum BRET signal. They also allow evaluation of BRET50, which is proposed to indicate the ability of two partners to interact (Audet et al., 2008); nonetheless, this would only be the case if the association between the receptors is reversible, which has not yet been demonstrated. Total fluorescence and luminescence are used as relative measures of total acceptor and donor protein expression, respectively. Total fluorescence is determined with an excitation filter at 485 nm and an emission filter at 535 nm. Total luminescence is measured 5 to 10 min after coelenterazine addition, in the absence of the emission filter. Because BRET-based experiments do not permit subcellular analysis, results can be altered, for example, by random collisions because of

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accumulation of chemokine receptors in intracellular membranes. To define receptor oligomers, experiments must be performed with several acceptor/donor ratios at a fixed surface density and a number of expression levels at a defined acceptor/donor ratio. Under these conditions, nonspecific interactions are independent of BRET efficiency and of acceptor/ donor ratios. Finally, it is very useful to include positive controls such as the donor genetically fused to the acceptor and, when possible, a known interacting receptor pair whose characteristics resemble those studied. Although the ideal negative control is a noninteracting protein similar to that analyzed, the acceptor protein is normally used alone.

3.2. Fluorescent resonance energy transfer (FRET) techniques In FRET, donor excitation energy is transferred to the acceptor by means of an induced dipole-dipole interaction; efficiency depends on the distance between and orientation of donor and acceptor fluorophores (Sekar and Periasami, 2003). Ideal dyes are photostable, have little intensity fluctuation, and are relatively small in size to minimize perturbation of the chemokine receptor. For all FRET methods, donor/acceptor choice is critical. Donor emission spectra should ideally have maximum overlap with acceptor absorption spectra, although acceptor and donor emissions should be clearly separable to minimize background interference. Fluorochrome incorporation into the protein must also be considered, because FRET sometimes requires the generation of chimeric constructs in which each chemokine receptor is fused to a different, modified form of green fluorescent protein (GFP). Some donor/acceptor combinations are blue (BFP)/GFP, CFP/YFP, green (GFP)/dsREd, GFP2/YFP, and YFP/dsRed. The most frequently used is, nonetheless, the CFP/YFP combination, because both are extremely bright, and this combination offers few technical problems (Pollok and Heim, 1999). BFP is a poor donor, because it is not especially bright, making FRET between BFP and GFP difficult to detect. dsRed is a poor acceptor, because it has a broad absorption spectrum and excites the same wavelength as the donor (GFP or YFP). FRET is sometimes evaluated on intact receptors with specific Ab conjugated to appropriately selected fluorescent dyes. Some common dyes are Alexa488, FITC, Cy3 or Cy2 as donor, and rhodamine-2, Alexa555, Cy3, or Cy5 as acceptor. Although Cy3 (donor) and Cy5 (acceptor) form a suitable fluorochrome pair, the Cy2 donor/Cy3 acceptor pair is often more convenient, because it allows use of the widely available 488-nm argon laser line. Secondary antibodies are occasionally needed, although the increased distance between fluorophores complicates FRET detection; this can be

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resolved with dye-labeled F(ab0 ) fragments. Because FRET depends on fluorochrome distance and orientation, low FRET efficiency values do not necessarily correlate with lack of dimerization. Several methods are used to determine and quantify FRET. The first is sensitized acceptor fluorescence, in which the donor fluorescent dye is excited and the acceptor signal is measured (Fig. 5.2). Another possibility is acceptor photobleaching, a method based on quenching donor fluorescence. Some donor photons are used to excite the acceptor, decreasing the emission energy detected. Photobleaching of the acceptor abolishes FRET, increasing donor light emission (Fig. 5.3). This method cannot be used for living cells, however, because exposure to extended laser energy is thought to damage the cell. Finally, fluorescence lifetime imaging microscopy (FLIM) measures a chromophore’s fluorescence lifetime, allowing spatial resolution of biochemical processes. The fluorescence lifetime of a donor dye decreases under FRET conditions, independently of fluorophore concentration (Periasami et al., 2002). To develop FRET assays by photobleaching with the CFP/YFP pair, plate HEK-293T cells (3.5
104 cells/ml) on poly-L-lysine–coated coverslips (20 mg/ml, 1 h, 37  C) and incubate (24 h, 37  C, 5% CO2). Cotransfect receptor combinations at a 1:1 ratio (one YFP- and one CFP-labeled receptor) and incubate (48 h, 37  C, 5% CO2); confirm equal levels of each receptor at the cell surface with standard flow cytometry analysis. Wash cells in PBS and fix with 4% formaldehyde (4 min, RT). Wash in A YFP emission

YFP

CFP

CFP emission B CFP

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Figure 5.2 Analysis of chemokine receptor dimerization by the sensitized acceptor fluorescence method for FRET. (A) Scheme illustrating the sensitized acceptor fluorescence method. CFP is excited with a 405-nm laser line and YFP emission detected at 530 nm. Alternately, a decrease in CFP emission can be detected at 460 to 500 nm. (B) Unstimulated HEK-293Tcells were transiently cotransfected with CCR2-CFP and CCR5-YFP, fixed, and FRETdetermined by detection of YFP emission. Images show CFP staining (left),YFP staining (middle), and FRET (right).

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Figure 5.3 Analysis of chemokine receptor dimerization by the photobleaching method in FRET. (A) Schematic representation of the photobleaching method. Some donor energy is used to excite the acceptor, decreasing detectable donor emission (left). Photobleaching of the acceptor abolishes FRET, increasing donor emission (right). (B) FRET evaluation of chemokine receptor heterodimerization with the acceptor photobleaching method. Unstimulated HEK293T cells were transiently cotransfected with a 1:1 combination of CCR2-CFP and CCR5-YFP, fixed, and FRET was determined. The image shows CFP staining before (CFP-pre) and after (CFP-post) photobleaching, as well as a false color merged image (FRET) and a zoom image of FRETat the photobleached area (insets).The DIC image is also included (left).

PBS and mount coverslips onto slides with PBS (pH 7.0) containing 80% glycerol. Evaluate fluorescence on a confocal microscope with appropriate filters for the fluorochrome. In a FRET assay, an image of the cell region of interest is taken with standard spectroscopic settings. CFP and YFP are excited by separate sweeps of the 405-nm (laser diodo [25 mW]) and 515-nm lines (three-line argon laser [45 mW]), respectively, and directed to the cell by means of a 405- to 440/515-nm dual dichroic mirror. The emitted fluorescence is split by a 510-nm dichroic mirror for CFP and directed to a spectral detector adjusted to the 460- to 500-nm range. For YFP, fluorescence is directed to a spectral detector adjusted to the 530- to 570-nm range. Confocal fluorescence intensity data (ICFPpre and IYFPpre) are recorded, with a pinhole of 100, as the average of four line scans per pixel and digitized at 12 bits. Repeated scans with 515 nm maximum light intensity are used to photobleach YFP, which requires 5 to 30 sec at maximal scan rates and a 100-pinhole aperture. After 60 to 90% of YFP bleaching, fluorescence intensity (ICFPpost and IYFPpost) is measured with identical parameters. With ImageJ 1.40g software (NIH), FRET efficiency is determined on a pixel-by-pixel basis (E) and calculated in percent as E = (1  FDpre/FDpost)
100%, where FDpre and FDpost are the background-corrected CFP fluorescence intensities before

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and after YFP photobleaching, respectively (Kenworthy, 2001; Zimmermann, 2002). As a negative control, FRET should be determined in transiently transfected HEK-293T cells with a chemokine receptor-CFP and in a region of the cotransfected cells without photobleaching. FRET efficiency should be calculated from several independent determinations, with at least 50 images from each. Recall that FRET provides meaningful data when determined with cells that have been evaluated individually for similar CFP and YFP fluorescence intensities. An alternative based on the sensitized acceptor fluorescence method allows FRET evaluation in cell populations (Carriba et al., 2008). HEK-293T cells are transiently transfected with vectors encoding the chemokine receptors of interest, coupled to CFP or GFP2 (donor) and YFP (acceptor). At 48 h posttransfection, distribute cell suspensions (20 mg protein/well) into 96-well microplates and read them in a fluorimeter equipped with a highenergy xenon flash lamp, with a 10-nm bandwidth excitation filter at 430 nm (CFP) or 400 nm (GFP2) and 10-nm bandwidth emission filters for 495 nm (CFP), 510 nm (GFP2), and 530 to 535 nm (YFP). To avoid spectral mixing, use identical gain settings for all experiments to maintain a constant relative contribution of the fluorophores to the detection channels. It is critical to measure the contribution of donor and acceptor alone to each detection channel in experiments with cells expressing only one of the proteins; these values are normalized to the sum of the signal obtained in the two detection channels. To exclude an effect on FRET efficiency because of the acceptor/ donor ratio and to be able to compare dimerization efficiency between receptor pairs, generate a FRET curve with constant expression levels of the receptor coupled to the donor and increasing amounts of the receptor coupled to the acceptor. This curve yields the FRETmax, which is not informative of interaction specificity, as well as the FRET50 (the acceptor/donor value at half-maximal FRET), which indicates the propensity of the interacting partners to associate and thus reflects differences in the relative affinity of these two partners. FRET efficiency is determined as previously (Zimmermann, 2002). FRET-based experiments can also be designed to determine the dynamics of receptor conformation at the cell membrane (Pello et al., 2008). If the role of the ligand is being studied, cells should be stimulated before they are fixed. The influence of a given receptor, R1, on R2:R2 homodimers can be determined by measuring photobleaching FRET in HEK-293T cells transiently cotransfected with CFP-R2 and YFP-R2 and comparing the result with FRET in HEK-293T cells transiently cotransfected with these two receptors plus R1. To facilitate analysis, R1 is expressed in a pIRES2AcGFP1-Nuc vector (Clontech). Only R1-expressing cells will be GFP-labeled in the nucleus and will be used to determine FRET. These experiments can also be modified to measure FRET between receptor heterodimers. In all cases, R1 expression in GFPþ cells should be controlled by flow cytometry and the CFP/YFP ratio determined by separate

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measurement of fluorescence levels. Controls should also include evaluation of the effect of R1 on R2 expression. For a complete analysis, the effect of the unlabeled receptor on FRET saturation curves assays should be measured; determine sensitized acceptor FRET in cells coexpressing a constant donor amount and increasing acceptor levels alone or in the presence of the unlabeled specific or unspecific receptors. FLIM is the most powerful FRET technique, because it is independent of transfection levels, although it requires complex equipment. In a typical FLIM determination, cells are plated on a chamber coverglass and transfected as described previously. Avoid fixing to allow in vivo measurements. FLIM is measured with a confocal microscope with a High Speed Lifetime Module and a 60
PlanApo 1.4 objective, or equivalent equipment. Fluorescence lifetime is determined after excitation with a pulsed laser (picosecond pulses) and a bandpass emission filter appropriate for the fluorochrome used and is quantitated with LIMO (Nikon) or similar software. Avoid the use of mounting solutions, which increase autofluorescence. The fluorescence lifetime of a donor is a constant parameter in specific experimental conditions (Fig. 5.4). A reduction in donor lifetime is due to a A

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Figure 5.4 Analysis of chemokine receptor dimerization by FLIM. (A) Scheme showing the FLIM method. The fluorescence lifetime of CFP after pulsed laser excitation (440 nm) (left) decreases under FRETconditions (right). (B) CFP fluorescence lifetime images (calculated from the phase shift) of HEK-293 cells expressing CCR5-CFP (left) or CCR5-CFP/CCR2-YFP (right). The pseudocolor scale ranges from 0 (black) to 4.0 nanosec (white).

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quenching effect triggered by the acceptor and is, therefore, indicative of chemokine receptor dimerization. Positive and negative controls as in BRET or in other FRET methods should also be included. Pay careful attention to variations in pH, temperature, and ionic strength in the medium, because these parameters cause alterations in FLIM measurements. Mycoplasma infection of the cells will also cause artifacts.

4. Sequential BRET-FRET (SRET) Technology Although BRET and FRET techniques are widely used to demonstrate homo- and heterodimers in living cells, they are inadequate for evaluating high-order complexes; that is, complexes involving more than two molecules. A new BRET-FRET–based technique called sequential BRET-FRET (SRET) was recently described (Carriba et al., 2008). SRET uses cells expressing a protein fused to RLuc, a protein fused to a BRET acceptor (GFP2 or YFP), and a protein fused to a FRET acceptor (YFP or DsRed). Addition of a RLuc substrate promotes acceptor excitation by BRET and subsequent energy transfer to the FRET acceptor (Fig. 5.5). We use suspensions of transiently cotransfected HEK-293T cells (20 mg protein/well) distributed in 96-well microplates, read in a fluorimeter equipped with a high-energy xenon flash lamp and a series of 10-nm bandwidth excitation and emission filters appropriate for the receptor-fused protein; this allows detection of BRET and FRET acceptor emission. These experiments require quantitation of receptor-fluorochrome expression, separation of the relative contribution of each fluorophore to the detection channels, quantitation of receptor-RLuc expression by determining luminescence, and, finally, SRET determination after addition of the RLuc substrate. The exact proportion of each fluorophore

RLuc 400 nm BRET

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Figure 5.5 Sequential resonance energy transfer technique (SRET). Cell coexpressing chemokine receptors, each fused to RLuc, to YFP, or to dsRED. Addition of the RLuc substrate (coelenterazine) triggers luciferase light emission at 485 nm. In consequence, the excited donor FRET (YFP) emits at 530 nm, which excites the FRET acceptor (dsRED). dsRED emission is then detected at 590 nm. This technique is used to study oligomerization.

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in the cell population is thus established, which indicates the receptor ratio used. These experiments measure FRET in cell populations, as for BRET; analyses, therefore, cannot be restricted to a specific cell area. All the general considerations and controls described for BRET and FRET are also applicable for these assays.

5. Conclusion Chemokines are the principal chemotactic factors implicated in the regulation of leukocyte traffic, both to inflammation sites and for establishing lymphoid organ architecture. Chemokines mediate their function by interacting with specific members of the seven-transmembrane, G-protein–coupled receptor family, which are expressed on the cell surface. Much information is available on the biochemical pathways activated by this large receptor family. Recent experiments, including those based on the application of resonance energy transfer (RET) technology, have, nonetheless, revealed an unexpected degree of complexity in chemokine receptor dynamics at the plasma membrane. In addition to the known promiscuity between ligands and receptors, the chemokine receptors can adopt a variety of conformations at the cell surface. These homo- and heterodimeric, and possibly oligomeric conformations, might be modulated by the levels of chemokine receptors or of other GPCR, as well as by chemokine expression. To explain the precise role of ligands and receptors in these dynamics, the classical biochemical techniques such as crosslinking, immunoprecipitation, and Western blot must be complemented by new technologies such as those based on RET and microscopy analysis. For these procedures, methodologic questions will need to be clarified, including use of different cell types, protein overexpression, and use of chemical inhibitors that can alter in vitro distribution, availability, and/or function of chemokine receptors compared with their in vivo behavior. Correctly used, these techniques will clearly be of value in unraveling the complexities of chemokines, their receptors, and their signals. Improved understanding of receptor homoand heterodimerization is changing our view of chemokine receptor structure and activation, which is likely to have substantial influence on drug development and screening.

ACKNOWLEDGMENTS We thank the members of the DIO chemokine group, who contributed to some of the work described in this review. We also thank C. Bastos and C. Mark for secretarial support and helpful editorial assistance, respectively. This work was partially funded by grants from the EU (Innochem LSHB-CT-2005-518167 and Molecular Imaging LSHG-CT-2003-503259),

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the Spanish Ministry of Science and Innovation (SAF2005-03388), and the Madrid Regional Government. The Department of Immunology and Oncology was founded and is supported by the Spanish National Research Council (CSIC) and by Pfizer.

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